Telomeres, A Busy Platform for Cell Signaling

Abstract

Telomeres are the terminal structures at the ends of linear chromosomes that represent a solution to the end replication problem. Specific binding of the six-protein subunit complex shelterin to telomeric, repetitive TTAGGG DNA sequences contributes to the stable architecture and maintenance of telomeres. Proteins involved in the DNA damage response are also localized at telomeres, and play a role in the surveillance and maintenance of telomere integrity. The enzyme responsible for telomere extension is telomerase, a ribonucleoprotein with reverse transcriptase activity. In the absence of telomerase, telomeres shorten to a length threshold that triggers the DNA damage response and replicative senescence. Here, we will summarize the latest findings concerning vertebrate telomere structure and epigenetics, and we present data regarding the impact of short telomeres upon cell signaling. In particular, in murine embryonic stem cells lacking telomerase, we found that distribution of cytosolic/nuclear β-catenin, a key component of the Wnt signaling pathway, changes when telomeres become critically short. We discuss implications and future perspectives of the effect of epigenetic modifications and/or conformational changes of telomeres on cell metabolism and signaling networks. Such an analysis may unveil potential therapeutic targets for pathologies like cancer, where the integrity of telomeres is altered.

Introduction

Telomere structure in vertebrates

Telomeres are the structures at the ends of chromosomes that protect them from end-to-end fusions and solve the problem of end replication, i.e., the loss of genetic material due to inherent limitations in the DNA replication process (Blackburn, 1991). Telomeres consists of a repeated six-nucleotide G-rich sequence, 5′-TTAGGG-3′, that is folded into a telomeric loop (t-loop) (Griffith et al., 1999). The telomere contains a double-stranded region and a single-stranded overhang, also referred to as the G-strand overhang, whose length is tightly regulated (Wright et al., 1997; Sfeir et al., 2005; Wu et al., 2012). Telomeres are protected and regulated by a specific hexaprotein complex, called shelterin (i.e., TRF1, TRF2, RAP1, TIN2, POT1, TPP1) (Figure 1A), and additional non-telomere specific proteins that are implicated in the cellular DNA damage response (de Lange, 2005; Longhese, 2008). Shelterin inhibits the ataxia telangiectasia mutated (ATM) and ATM and Rad3-related (ATR)-dependent DNA damage response, non-homologous end joining and homologous recombination DNA repair pathways, and resection by 5′-exonucleases (Sfeir and de Lange, 2012). Some of these activities are specific to shelterin whereas other activities that inhibit non-homologous end joining and resection are supported by other telomere-associated proteins such as Ku70/80 and 53BP1, respectively (Sfeir and de Lange, 2012). As the enzyme responsible for telomere extension, telomerase is a key factor that contributes to chromosome end protection (Blackburn et al., 1989). Telomerase is a reverse transcriptase that copies a stably associated RNA template into telomere DNA (Greider and Blackburn, 1987, 1989; Shippen-Lentz and Blackburn, 1990). In mice, the extension of telomeres occurs during S-phase and telomerase extends the shortest telomeres preferentially (Hemann and Greider, 1999; Samper et al., 2001; Erdmann et al., 2004; Stern and Bryan, 2008).

Figure 1

Telomere epigenetics

Telomeric DNA contains nucleosomes, although the nature of telomeric chromatin is peculiar (Makarov et al., 1993) (Figure 1B). Simplistically, nucleosomes and shelterin compete with each other for the binding of telomeric DNA, hence it is not surprising that TRF2 influences the positioning of the nucleosomes, i.e., the nucleosome abundance at telomeres is inversely correlated with the amount of TRF2 (Benetti et al., 2008b; Galati et al., 2012). Nucleosome spacing by TRF2 occurs in S/G2 phase, which coincides with the end of DNA replication and telomere replication (Galati et al., 2012). Epigenetic marks, notably histone and DNA methylation, in sub-telomeric and telomeric regions contribute further to telomere maintenance and stability (Blasco, 2007) (Figure 1B). A high degree of DNA methylation guarantees a closed chromatin state that is associated with gene silencing in regions upstream of the telomeres. This phenomenon, first described in Drosophila melanogaster and yeast, is known as the telomere positioning effect or TPE (Levis et al., 1985; Gottschling et al., 1990; Nimmo et al., 1994). In humans, telomere length positively affects TPE through a change in the conformation of chromatin (Baur et al., 2001). Epigenetic defects at telomeres, such as those driven by the loss of DNA methyl transferases or histone methyl transferases, lead to telomere defects that result in aberrant telomere lengthening attributed partially to an increase in homologous recombination (Gonzalo et al., 2006; Benetti et al., 2007b, 2008a). In mice lacking the telomerase RNA component (mTerc^−/−), short telomeres are associated with epigenetic changes at the telomeres, i.e., a decrease of tri-methylated histone 3 and histone 4 and an increase in histone acetylation (Benetti et al., 2007a). Thus, critically shortened telomeres show signs of an “open” chromatin state that favors recombination events (Benetti et al., 2008a). It is reasonable to postulate that epigenetic changes in telomeric DNA and histones affect the binding of shelterin and, in turn, affect telomere structure and the recruitment of telomerase (Blasco, 2007).

The complex regulation of telomeric structure, maintenance, and epigenetics has been underscored further by the discovery of the transcription of telomeres into a telomere repeat-containing RNA (TERRA) that contains UUAGGG repeats (Azzalin et al., 2007; Schoeftner and Blasco, 2008) (Figure 1B). The length and amount of TERRAs are directly correlated with telomere length and vary with the cell cycle. Because of their ability to anneal with the template sequence in the telomerase RNA component, TERRAs are able to inhibit telomerase (Figure 1B) (Redon et al., 2010). The precise role of TERRAs has not yet been established fully, but TERRAs are proving to be an interesting regulator of telomere dynamics.

Telomeres, telomerase, and the Wnt signaling pathway

Telomere dysfunction is also linked to perturbation of other cellular processes that include the Wnt/β-catenin signaling network. The Wnt/β-catenin signaling cascade controls many aspects of organism development, cell proliferation, and differentiation (Valenta et al., 2012). In the absence of Wnt, β-catenin is phosphorylated and rapidly degraded by a destruction complex containing Axin, APC, CK1, and GSK3β (Clevers and Nusse, 2012). However, in the presence of Wnt, β-catenin is stabilized and imported into the nucleus where, together with the transcription complex TCF/LEF, it regulates the transcription of Wnt target genes (Behrens et al., 1996; Molenaar et al., 1996). Cytoplasmic β-catenin localizes to the cell membrane through an interaction with E-cadherin and serves to stabilize cell adhesion (Ozawa et al., 1989).

The first link between telomerase and Wnt signaling was suggested from an analysis of transcription profiles of mouse and human cells expressing catalytically active or inactive Tert (Choi et al., 2008). Stem cells that express mTert, irrespective of its competence for catalytic activity, exhibit transcriptional activation of genes regulated by Wnt (Choi et al., 2008). In addition, mTert is localized to the promoters of genes regulated by Wnt3a and β-catenin (Park et al., 2009). In mESC over-expressing mTert, the activation of Wnt signaling by LiCl leads to the transcriptional activation of β-catenin (Park et al., 2009). However, another study compared the transcriptional profile of cells from mTert^−/− mice with mTerc^−/− mice, and observed no substantial difference in gene expression (Vidal-Cardenas and Greider, 2010). In particular, the Wnt signaling network was unaffected, and the authors suggested that the link between telomerase and Wnt signaling might be a neomorph due to telomerase over-expression (Strong et al., 2011). More recently, it has been found that β-catenin can regulate mTert transcription in mESC (Hoffmeyer et al., 2012). This regulation involves Klf4, one of the four transcription factors required to induce pluripotent stem cells. The control operated on mTert by β-catenin may be direct because β-catenin occupies the mTert promoter (Hoffmeyer et al., 2012). β-catenin also activates TRF2 transcription (Diala et al., 2013). Finally, c-myc, which is also under the control of β-catenin/Wnt signaling, is a known regulator of mTert transcription (Wang et al., 1998), thus implying a very tight regulation of this gene and the involvement of multiple signaling networks (Greider, 2012).

Telomere attrition triggers activation of the DNA damage response and other changes that herald the onset of genome instability (Cimprich and Cortez, 2008; Schoeftner and Blasco, 2010). To dissect the complexity of such processes, it is important to distinguish between the impact of telomerase loss versus the impact on telomere length. In this regard, murine embryonic stem cells represent a valuable model system and in ESC lacking mTert, we show that critically short telomeres (and not telomerase presence per se) can impact cell signaling cascades even in the cytoplasm. We focused on β-catenin because of its known link to telomere function and because its dynamic phosphorylation-dependent regulation appeared a logical choice for a first examination of the impact of DNA damage signaling at the telomere in the cytoplasm. Our data suggest that alteration of telomere structure or epigenetic modifications elicited by telomere shortening impacts cell signaling in extra-nuclear locations, which in turn may affect cell adhesion, metabolism, and protein turnover.

Results

Short telomeres affect cell adhesion and β-catenin distribution

Murine ESC lacking the telomerase reverse transcriptase were generated and characterized previously and show an accumulation of telomere signal-free ends at late passage (Liu et al., 2000; Erdmann et al., 2004). We queried whether the abundance of key signaling factors would be altered in the presence of short telomeres, and focused our investigation on β-catenin, a critical component of the Wnt signaling network that controls cell proliferation and differentiation (Clevers and Nusse, 2012). β-catenin distribution and post-translational modifications were compared in mTert^−/− at late passage (>60 passages) and wild-type ESC at a similar passage number (Figure 2A). We observed that cytosolic β-catenin was significantly more abundant in mTert^−/− ESC with critically short telomeres compared to wild-type cells (Figure 2B, Student’s t-test P = 0.003) while the total content remained unchanged (Figure 2B, P = 0.968). Accordingly, higher levels of nuclear β-catenin were observed in wild-type cells (Figure 2C, P = 0.027). Taken together, these results indicate that the distribution of β-catenin differed between the two cell types.

Figure 2

β-Catenin is a target of the GSK3β kinase which phosphorylates the residues S33/37, and T41. The tri-phosphorylated form of β-catenin is rapidly degraded by the proteasome (Liu et al., 2002). We used an antibody specific for the triple-phosphorylated β-catenin (S33/37, T41) to assess the phosphorylation status of β-catenin in ESCs with or without short telomeres, and found no difference in the levels of phosphorylated, cytosolic β-catenin (Figure 2E). Since the degradation of phospho-β-catenin occurs very rapidly and may mask subtle differences in abundance, we treated ESCs with the proteasome inhibitor MG132. In the presence of MG132, the difference in the phosphorylation status of cytosolic β-catenin in wild-type cells compared to mTert^−/− ESC achieved statistical significance (Figure 2D, P = 0.0014). These results suggest that β-catenin is degraded less rapidly in mTert^−/− ESC with short telomeres, or that there is a pool of β-catenin in cells with critically short telomeres that is immune to proteasome-dependent degradation.

As GSK3β activity is inhibited by the phosphorylation of a serine at amino acid position 9 (Sutherland et al., 1993; Desbois-Mouthon et al., 2001; Fukumoto et al., 2001), we assessed the serine 9 phosphorylation status of GSK3β. We did not detect a significant difference between wild-type and mTert^−/− ESCs (Figure 2E). GSK3β is phosphorylated by the kinase AKT, whose activity is regulated by the phosphorylation of serine 473 (Alessi et al., 1997; Fukumoto et al., 2001). We did not observe a significant difference in the level of AKT phosphorylation between wild-type and mTert^−/− cells (Figure 2E). These results suggest that downstream effectors of Wnt signaling remain unaltered in mTert^−/− cells with critically short telomeres.

Comparison of Wnt signal transduction

LiCl is an inhibitor of GSK3β that triggers phosphorylation on serine 9 through an as yet unknown mechanism (Rao et al., 2005a). Because inactivation of the kinase activity of GSK3β results in the inhibition of phosphorylation of β-catenin and its stabilization, LiCl treatment is often used to activate Wnt (Rao et al., 2005b). To assess whether the different distribution of β-catenin was associated with a difference in Wnt signaling, we treated mESC with LiCl and, as expected, observed a stabilization of β-catenin levels in both wild-type and mTert^−/− ESCs (Figure 3A). We did not detect a significant difference in the transcription of a specific target of Wnt signaling, Axin2, in response to Wnt3a (Figure 3B). Similar to Axin2, a reporter system containing three consensus TCF binding sites upstream of the firefly luciferase gene did not exhibit a statistically significant difference between WT and KO cells (Figure 3C) (Korinek et al., 1998). Thus, two independent outputs of Wnt signaling were not appreciably altered in mTert^−/− ESCs with short telomeres. The transcription of the cell cycle-regulated genes c-myc and cyclinD1 are also regulated by Wnt and many other signaling networks, but did not exhibit a statistically significant trend in response to Wnt3a (Burdon et al., 2002; Jho et al., 2002) (data not shown).

Figure 3

In order to identify factors responsible for β-catenin cytosolic accumulation in mTert^−/− ESCs with short telomeres, we compared the profile of β-catenin interacting proteins using mass spectrometry. Three independent β-catenin immunoprecipitations from total lysates were performed and only proteins recovered in all three experiments were considered (152 proteins in total). We considered a protein interaction significantly different between wild-type or mTert^−/− ESC if it exhibited a peptide intensity ratio of<0.667 or>1.5 with an unpaired Student’s t-test P value < 0.05 (see Section Materials and Methods). Adenomatous polyposis coli (APC), was enriched by approximately twofold, (P = 0.03) in mTert^−/− ESC relative to wild-type ESC (Table 1). APC interacts with β-catenin and together with Axin1, constitutes the scaffold of the destruction complex that regulates the stability of cytosolic β-catenin (Rubinfeld et al., 1993; Hart et al., 1998; Hamada and Bienz, 2004; Clevers and Nusse, 2012). Interestingly APC is also implicated in the regulation of the nuclear export of β-catenin and, therefore, influence the balance between nuclear and cytoplasmic β-catenin independently of Wnt signaling (Henderson, 2000). Eight other proteins were also identified as novel interactors of β-catenin and have not yet been further characterized (Table 1).

Rescue of cytosolic β-catenin with telomere lengthening

To address whether the reintroduction of telomerase and extension of telomeres could restore the level of cytosolic β-catenin, we reintroduced mTert into mTert^−/− ESCs under the control of a tetracycline-inducible promoter and, after selection of mTert-positive clones, cells were propagated under mTert induction conditions (+Dox) for 70 days (Figures 4A,B). The reactivation of telomerase upon addition of doxycycline was confirmed by TRAP (telomerase repeat amplification protocol, data not shown) and the extension of the telomeres verified by Q-FISH analysis (Figure 4B). At this point, the culture was split in two and propagated in the absence (−Dox) or presence (+Dox) of mTert for an additional four population doublings (Figures 4A,B). Analysis of the level of cytosolic β-catenin in ESCs with extended telomeres, irrespective of mTert expression, revealed a rescue of the cytosolic β-catenin to levels comparable to wild-type ESCs (Figure 4C, P = 0.20). This result suggests that the distribution of β-catenin is dependent on telomere length rather than telomerase activity. This result is similar to the finding that mice or ESCs lacking telomerase activity do not exhibit phenotypes until telomeres become critically shortened (Erdmann and Harrington, 2009; Strong et al., 2011). Instead, a loss of tissue self-renewal is evident at generations above G4, underscoring the dependence of the phenotype upon loss of telomere integrity (Vidal-Cardenas and Greider, 2010; Strong et al., 2011).

Figure 4

Discussion

Here, we discussed the impact of telomere integrity on cell signaling. We show new data that mESC with short telomeres undergo an accumulation of cytosolic β-catenin. Although the level of nuclear β-catenin is higher in wild-type cells, this difference does not result in an induction in the transcription of Wnt target genes. This observation is in general agreement with the finding that activation of Wnt signaling leads to β-catenin nuclear import, but there is no relationship between the level of nuclear β-catenin and Wnt activation (Guger and Gumbiner, 2000). The higher cytosolic content of β-catenin in mTert^−/− ESCs might be the result of an altered balance of β-catenin nuclear import/export or of β-catenin degradation/stabilization. In support of the first explanation, we observed an enrichment of APC in β-catenin immunoprecipitates from mTert^−/− cells. APC shuttles between the nucleus and the cytoplasm independently from other factors of the destruction complex (Henderson, 2000). The destruction complex is not disassembled in the presence of Wnt; instead, degradation of β-catenin by the proteasome is altered upon Wnt stimulation (Hilger and Mann, 2012; Li et al., 2012). This finding may explain why an increased level of APC-β-catenin complex might not necessarily result in higher β-catenin degradation. Furthermore, our finding that the phosphorylation of β-catenin is increased in wild-type ESCs but not mTert^−/− ESCs in the presence of the proteasome inhibitor MG132 supports the notion that the activity of the proteasome is altered in mTert^−/− ESCs with critically short telomeres. Such an effect on the proteasome may not be surprising as previous studies have demonstrated a link between cellular aging and an alteration of the ubiquitin-proteasome machinery (Grillari et al., 2006). These results support the notion that the up-regulation of β-catenin occurs as result of altered protein degradation in the presence of short telomeres. Further analysis of the complex composition, stoichiometry, and relative abundance of these complexes in the nucleus and cytoplasm in response to critically short telomeres will be informative.

It remains to be tested whether difference in β-catenin phosphorylation status in murine ESCs with critically short telomeres also impacts Wnt signaling more generally. β-catenin intersects several signaling cascades, not all of which are linked to Wnt (Valenta et al., 2012). For example, in the presence of LIF, β-catenin transcriptional activity is not required to maintain self-renewal; it is mostly its role at the cell adhesion structures that is required to allow differentiation (Lyashenko et al., 2011). On the cytoplasmic side of the plasma membrane, β-catenin interacts with cadherins and α-catenin to stabilize cell–cell adhesion structures but also to regulate cytoskeleton dynamics (Yamada et al., 2005). The majority of these interactions with its partners are regulated by phosphorylations at sites other than the S33, 37, T41 (Liu, 1999). In general, the pattern of β-catenin phosphorylation regulates the transition from a structural versus signaling role (Valenta et al., 2012). Thus, the impact of critically short telomeres upon the phophorylation of β-catenin at sites other than S33, 37, T41 should also be investigated.

In conclusion, global genomic changes driven by short telomeres have consequences on general gene expression and cell metabolism (Figures 5A,B). For example, critically short telomeres in mESCs lacking Tert influence DNA and histone methylation at the promoters of pluripotency regulators such as Nanog and Oct4, thereby negatively affecting the stable differentiation of mESCs (Pucci et al., 2013). In the nuclear compartment, β-catenin interacts with several chromatin remodeling complexes, including the histone acetylase p300/CBP and the helicases TIP49a/Pontin52 and TIP49b/Pontin52 and Brg1, the latter of which also interacts with telomerase (Mosimann et al., 2009; Park et al., 2009). β-catenin increases H3K4 methylation at the c-myc promoter through its interaction with the histone methyltransferase SET1. Interestingly, this activity is counteracted by Apc, which displaces β-catenin from the chromatin remodeling complex (Sierra et al., 2006). Furthermore, the involvement of β-catenin in the control of telomerase transcription and the previous finding of complexes containing both telomerase and β-catenin at promoter sequences show that β-catenin and telomere integrity are connected (Park et al., 2009; Hoffmeyer et al., 2012). Hence, it is reasonable to postulate that the alteration of binding of telomeric proteins or epigenetic modifications can trigger signaling cascades that might culminate in changes at the plasma membrane and alter communication with the environment (Figures 5A,B). Although the precise means by which short telomeres elicit genome-wide changes in gene expression is unknown, one candidate mechanism is RAP1, a transcription factor that binds extra-telomeric sites and in whose absence there are a number of changes in gene expression in processes related to cell metabolism, cell adhesion, and cancer (Martinez et al., 2010; Martinez and Blasco, 2011). Moreover, together with Trf2, Rap1 transcription is directly regulated by β-catenin (Diala et al., 2013). Taken together, these findings reinforce the notion that β-catenin and telomere structure and function are interconnected. Clearly, the future promises to uncover additional intriguing links between the impact of critically short telomeres and cytoplasmic cell signaling.

Figure 5

Materials and Methods

Cell culture

Wild-type and telomerase reverse transcriptase-deficient mESC (E14) were cultured in GMEM, 15% v/v FBS (Hyclone, UK), β-mercaptoethanol, penicillin/streptomycin, and leukemia inhibitory factor (Sigma, UK), and split by a ratio of 1:8 every 3 days, as described in Erdmann et al. (2004). The absence or presence of telomerase activity was assessed by the TRAP (telomerase repeat amplification protocol), performed following manufacturer’s instructions (TRAPeze, Millipore, UK). Cells grown in 6-well plates were lysed in 50 μL of CHAPS 1 × buffer. Two microliters of cell lysate were assayed in the TRAP.

Cell fractionation and immunoblotting

Cells grown in 10-cm diameter plates were washed and scraped in PBS. For cell fractionation, cells were pelleted for 5 min at 1500 × g and re-suspended in hypotonic lysis buffer (50 mM Tris, pH 7.8, 250 mM sucrose, 2 mM EDTA) supplemented with Roche’s Complete Protease Inhibitor Cocktail and PhoSTOP Phosphatase Inhibitor Cocktail. Cells were homogenized with 20 strokes in a Dounce homogenizer and then centrifuged for 10 min at 2800 × g to precipitate the nuclei. The supernatant represented the cytosolic fraction. The nuclear pellet was re-suspended in buffer S1 (0.25 M sucrose, 10 mM MgCl₂), layered over an equal volume of buffer S3 (0.88 M sucrose, 0.5 M MgCl₂), and centrifuged at 2800 × g for 10 min. The pellet was re-suspended in RIPA buffer (50 mM Tris, pH 7.5, 150 mM NaCl, 1% v/v NP-40, 0.5% w/v deoxycholic acid) containing protease and phosphatase inhibitors. The nuclear extract was sonicated and centrifuged at maximum speed 16,000 × g for 10 min to obtain the nuclear extract. The protein concentrations of the cytosolic and nuclear lysates were measured using the Bradford method prior to loading onto gels for SDS-PAGE. In all experiments, 10 μg of cytosolic protein and 2 μg of nuclear protein were loaded. For total cell extracts, cells were scraped and re-suspended in RIPA buffer. Ten micrograms of total protein lysate, measured by the Bradford method, were loaded onto gels for SDS-PAGE. SDS-PAGE was performed with NuPAGE Bis–Tris 4–12% w/v gradient gels (Invitrogen, UK). After electrophoresis, the proteins were transferred onto an Immobilon-FL membrane (Millipore) at a constant 100 V for 1 h. The membrane was blocked in 5% w/v skimmed milk powder (non-fat) in TBST. Primary antibodies were incubated with the membrane overnight at 4 °C in 2.5% w/v skimmed milk powder in TBST. The primary antibodies used were as follows: rabbit polyclonal anti-β-catenin (1:4000; Bethyl Laboratories, Inc., USA); rabbit monoclonal anti-β-catenin, clone E247 (1:4000; Millipore); rabbit polyclonal anti-phospho-β-catenin (Ser33/37/Thr41) (1:2000; Cell Signaling UK); anti-E-cadherin (1:4000; BD Biosciences UK); rabbit anti-phospho-GSK3β (S9) and mouse anti-GSK3β total (both at 1:1000; Cell Signaling); rabbit anti-phospho-AKT (S473) and rabbit anti-AKT total (both at 1:2000; Cell Signaling). Mouse anti-β-tubulin (1:4000; Sigma) was used as a loading control for the total protein extracts and cytosolic fractions, whereas rabbit anti-lamin B1 (1:2000; a gift from Dr. Eric Schirmer) was used for nuclear fractions. Secondary antibodies were HRP-conjugated anti-mouse and anti-rabbit (1:10000 and 1:5000, respectively; GE Healthcare). For quantification of the immunoblot bands, secondary antibodies were donkey anti-mouse (IRDye 800) and donkey anti-rabbit (IRDye 680) (1:10000 and 1:5000, respectively; LI-COR Biosciences, UK). For Wnt3a treatment, 100 ng/mL of recombinant mouse Wnt3a (Millipore) was added to the culture medium for 3 days. For LiCl treatment, 30 mM LiCl was added to the culture medium for 4 h. For the inhibition of the proteasome, cells were treated with 10 μM of MG132 (Sigma, UK, dissolved in DMSO) for 6 h. Cells were collected by scraping and lysed as previously described for cell fractionation and western blot analysis.

Western blot quantitative analysis

Images of membranes probed with secondary IRDye antibodies were acquired with an Odyssey scanner and analyzed with Odyssey software (Licor Biosciences). Excel and GraphPad Prism v.5 were used for statistical analysis. Briefly, two rectangles of the same size were placed over β-catenin and the relevant control (β-tubulin or lamin B1 for cytosol or nuclear extract, respectively). The intensity of β-catenin was normalized to the value of the loading control within the same lane and averaged against at least three independent replicates. The Student’s t-test was used to evaluate the statistical significance of the comparison (Gardano et al., 2011).

Quantitative fluorescence in situ hybridization

The Q-FISH protocol was carried out as described (Liu et al., 2000). Metaphase spreads were captured using Metafer 4 software and analyzed using Isis software. Statistical analysis of telomere intensity distribution was performed using Welch’s unpaired t-test.

qRT-PCR

RNA was extracted from cells grown in 6-well plates using Qiagen’s RNeasy Mini Kit. The RNA was treated with DNase for 1 h prior to the reverse transcription reaction. One microgram of RNA was retrotranscribed with random primers (Invitrogen) using SMART MMLV reverse transcriptase (Clontech Laboratories, Inc., USA). The cDNA mixture was diluted 20 times in water containing RNase before proceeding with the qPCR (Lightcycler 480, Roche, UK). The sequences of the primers used were as follows: Axin2 forward 5′-AGCGCCAACGACAGCGAGTT-3′; Axin2 reverse 5′-TCCCCATGCGGTAAGGAGGGAC-3′; GAPDH forward 5′-AGGTCGGTGTGAACGGATTTG-3′; GAPDH reverse 5′-TGTAGACCATGTAGTTGAGGTCA-3′mTERT forward 5′-TTCTAGACTTGCAGGTGAACAGCC-3′; mTERT reverse 5′-TTCCTAACACGCTGGTCAAAGGGA-3′. Data were analyzed with Excel and GraphPad Prism v.5.

Top-flash experiments

Cells were seeded at a concentration of 2.5 × 10⁴ mL in 12-well dishes and, 24 h later, Extreme Gene 9 (Roche, UK) was used to transfect 0.5 μg DNA (in total) consisting of Top-firefly luciferase plasmid or the negative control Fop-firefly luciferase (Millipore) and 0.05 μg of Renilla plasmid transcribed with a SV40 promoter, pRL (Promega). Cell lysis was performed 48 h after transfection with the Passive Lysis buffer supplied by the Dual luciferase assay (Promega, UK). Firefly and Renilla luciferase activities were monitored following manufacturer’s instructions. Luciferase activity was recorded using an Infinite 200 instrument (Tecan group Ltd.). Wnt3a treatment was performed as previously described. Top-firefly luciferase signals were normalized to renilla luciferase values and then normalized to Fop-luciferase activity for each respective treatment, i.e., with or without Wnt3a. Graphpad Prism v.5 was used for statistical analysis.

mTERT cell transfection and plasmids

The plasmid pTRE-Bi-Tert-IRES-EGFP-Hygro was constructed by amplification of Tert cDNA by PCR and insertion into pTRE-Tight-Bi (Clontech) following digestion with EcoRI and SalI. IRES-EGFP sequence was obtained from pCAGMKOSiE (kindly provided by K. Kaji) and inserted into pTRE-Tight-Bi (following digestion with SalI and EcoRV) using SalI and HpaI sites and then inserted into pTRE-Bi-Tert using NotI sites. Finally, the hygromycin resistance gene was cloned by PCR into the XbaI restriction site of pTRE-Tight-Bi and pTRE-Bi-Tert-IRES-EGFP vectors to create pTRE-Bi-EGFP-Hygro and pTRE-Bi-Tert-IRES-EGFPHygro. The pCAG-rtTA-advanced vector was constructed by removal of the MKOS ORFs from CAGMKOSiE with EcoRI and BamHI and replacement with the advanced tetracycline reverse transactivator sequence (Clontech) (Pucci et al., 2013).

Immunoprecipitation and mass spectrometry

Wild-type and mTert^−/− ESC were propagated in three 15-cm diameter plates for each immunoprecipitation. Three independent immunoprecipitations were performed contemporaneously on wild-type and mTert^−/− ESC. Cells were lysed in 1 mL of lysis buffer (50 mM Tris, pH 7.5, 5 mM EDTA, 5 mM NaF, 10% v/v glycerol, 0.1% v/v NP-40, 1 mM DTT) supplemented with Roche’s Complete Protease Inhibitor Cocktail. Following centrifugation at 16,000 × g for 10 min, the amount of total protein in all the samples was assessed by the Bradford method. Ten micrograms of rabbit monoclonal anti-β-catenin antibody (clone E247, Millipore) were added to each lysate and incubated for 2 h at 4 °C with rocking. Magnetic Dynabeads Protein A (Invitrogen) was equilibrated in the lysis buffer prior to addition to cell lysates (10 μL beads added to each immunoprecipitation). The bead/lysate mixtures were then incubated for 40 min at 4 °C. Following four washes with a washing buffer (lysis buffer without glycerol), the beads were re-suspended in 20 μL washing buffer. The samples were boiled in Laemmli buffer and loaded onto a NuPAGE Bis–Tris 4–12% v/v gradient gel for SDS-PAGE. The gel was stained with SimplyBlue SafeStain (Life Technologies, UK). Each entire gel lane was sliced into six pieces, then processed according to an in-gel protocol for trypsin digestion.

Capillary-HPLC-MS/MS analysis was performed on an on-line system consisting of a micro-pump (1200 binary HPLC system, Agilent, UK) coupled to a hybrid LTQ-Orbitrap XL instrument (Thermo-Fisher, UK). MS/MS data was searched using MASCOT (Matrix Science Ltd, UK) against the Mus musculus subset of the NCBI protein database using a maximum missed-cut value of 2. Variable methionine oxidation, ST and Y phosphorylation, and N-term acetylation were used and fixed cysteine carbamidomethylation were used in all searches; precursor mass tolerance was set to 7 ppm and MS/MS tolerance to 0.4 amu. The significance threshold (p) was set below 0.05 (MudPIT scoring). A peptide Mascot score cut-off of 20 was used in the final analysis, which corresponds to a global false discovery rate of 3.6% using a decoy database search. LC-MS label-free quantification was performed using Progenesis (Non-linear Dynamics, UK). For label-free quantitation, the total number of Features (i.e., intensity signal at a given retention time and m/z) was reduced to MS/MS peaks with charge of 2, 3, or 4+ and only the five most intense MS/MS spectra were retained per “Feature.” The subset of multicharged ions (2+, 3+, 4+) was extracted from each LC-MS run. Protein quantification was performed as follows; for each protein, the associated unique peptide ions were summed to generate an abundance value and normalized by dividing the protein intensity by the bait intensity (β-catenin). The within group means were calculated to determine the fold change and a t-test was used between the two groups. Regarding quantitative cut-off thresholds, proteins were considered a hit if two or more peptides were detected with an absolute ratio of at least 1.5 (i.e., 1.5 fold increase, or 0.667 decrease) and a significance of p < 0.05. Nine proteins met this threshold criteria (Table 1).

References

• 1

  AlessiD. R.JamesS. R.DownesC. P.HolmesA. B.GaffneyP. R.ReeseC. B.et al (1997). Characterization of a 3-phosphoinositide-dependent protein kinase which phosphorylates and activates protein kinase Balpha. Curr. Biol.7, 261–269. 10.1016/S0960-9822(06)00122-9

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 2

  AzzalinC. M.ReichenbachP.KhoriauliL.GiulottoE.LingnerJ. (2007). Telomeric repeat containing RNA and RNA surveillance factors at mammalian chromosome ends. Science318, 798–801. 10.1126/science.1147182

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 3

  BaurJ. A.ZouY.ShayJ. W.WrightW. E. (2001). Telomere position effect in human cells. Science292, 2075–2077. 10.1126/science.1062329

  • CrossRef
  • Google Scholar

• 4

  BehrensJ.Von KriesJ. P.KuhlM.BruhnL.WedlichD.GrosschedlR.et al (1996). Functional interaction of beta-catenin with the transcription factor LEF-1. Nature382, 638–642. 10.1038/382638a0

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 5

  BenettiR.Garcia-CaoM.BlascoM. A. (2007a). Telomere length regulates the epigenetic status of mammalian telomeres and subtelomeres. Nat. Genet.39, 243–250. 10.1038/ng1952

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 6

  BenettiR.GonzaloS.JacoI.SchottaG.KlattP.JenuweinT.et al (2007b). Suv4-20h deficiency results in telomere elongation and derepression of telomere recombination. J. Cell Biol.178, 925–936. 10.1083/jcb.200703081

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 7

  BenettiR.GonzaloS.JacoI.MunozP.GonzalezS.SchoeftnerS.et al (2008a). A mammalian microRNA cluster controls DNA methylation and telomere recombination via Rbl2-dependent regulation of DNA methyltransferases. Nat. Struct. Mol. Biol.15, 998. 10.1038/nsmb.1399

  • CrossRef
  • Google Scholar

• 8

  BenettiR.SchoeftnerS.MunozP.BlascoM. A. (2008b). Role of TRF2 in the assembly of telomeric chromatin. Cell Cycle7, 3461–3468. 10.4161/cc.7.21.7013

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 9

  BlackburnE. H. (1991). Structure and function of telomeres. Nature350, 569–573. 10.1038/350569a0

  • CrossRef
  • Google Scholar

• 10

  BlackburnE. H.GreiderC. W.HendersonE.LeeM. S.ShampayJ.Shippen-LentzD. (1989). Recognition and elongation of telomeres by telomerase. Genome31, 553–560. 10.1139/g89-104

  • CrossRef
  • Google Scholar

• 11

  BlascoM. A. (2007). The epigenetic regulation of mammalian telomeres. Nat. Rev. Genet.8, 299–309. 10.1038/nrg2047

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 12

  BurdonT.SmithA.SavatierP. (2002). Signalling, cell cycle and pluripotency in embryonic stem cells. Trends Cell Biol.12, 432–438. 10.1016/S0962-8924(02)02352-8

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 13

  ChoiJ.SouthworthL. K.SarinK. Y.VenteicherA. S.MaW.ChangW.et al (2008). TERT promotes epithelial proliferation through transcriptional control of a Myc- and Wnt-related developmental program. PLoS Genet.4:e10. 10.1371/journal.pgen.0040010

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 14

  CimprichK. A.CortezD. (2008). ATR: an essential regulator of genome integrity. Nat. Rev. Mol. Cell Biol.9, 616–627. 10.1038/nrm2450

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 15

  CleversH.NusseR. (2012). Wnt/beta-catenin signaling and disease. Cell149, 1192–1205. 10.1016/j.cell.2012.05.012

  • CrossRef
  • Google Scholar

• 16

  de LangeT. (2005). Shelterin: the protein complex that shapes and safeguards human telomeres. Genes Dev.19, 2100–2110. 10.1101/gad.1346005

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 17

  Desbois-MouthonC.CadoretA.Blivet-Van EggelpoelM. J.BertrandF.CherquiG.PerretC.et al (2001). Insulin and IGF-1 stimulate the beta-catenin pathway through two signalling cascades involving GSK-3beta inhibition and Ras activation. Oncogene20, 252–259. 10.1038/sj.onc.1204064

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 18

  DialaI.WagnerN.MagdinierF.ShkreliM.SirakovM.BauwensS.et al (2013). Telomere protection and TRF2 expression are enhanced by the canonical Wnt signalling pathway. EMBO Rep.14, 356–363. 10.1038/embor.2013.16

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 19

  ErdmannN.HarringtonL. A. (2009). No attenuation of the ATM-dependent DNA damage response in murine telomerase-deficient cells. DNA Repair (Amst.)8, 347–353. 10.1016/j.dnarep.2008.11.009

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 20

  ErdmannN.LiuY.HarringtonL. (2004). Distinct dosage requirements for the maintenance of long and short telomeres in mTert heterozygous mice. Proc. Natl. Acad. Sci. U.S.A.101, 6080–6085. 10.1073/pnas.0401580101

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 21

  FukumotoS.HsiehC. M.MaemuraK.LayneM. D.YetS. F.LeeK. H.et al (2001). Akt participation in the Wnt signaling pathway through Dishevelled. J. Biol. Chem.276, 17479–17483. 10.1074/jbc.C000880200

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 22

  GalatiA.MagdinierF.ColasantiV.BauwensS.PinteS.RicordyR.et al (2012). TRF2 controls telomeric nucleosome organization in a cell cycle phase-dependent manner. PLoS ONE7:e34386. 10.1371/journal.pone.0034386

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 23

  GardanoL.HollandL.OultonR.Le BihanT.HarringtonL. (2011). Native gel electrophoresis of human telomerase distinguishes active complexes with or without dyskerin. Nucleic Acids Res.40, e36. 10.1093/nar/gkr1243

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 24

  GonzaloS.JacoI.FragaM. F.ChenT.LiE.EstellerM.et al (2006). DNA methyltransferases control telomere length and telomere recombination in mammalian cells. Nat. Cell Biol.8, 416–424. 10.1038/ncb1386

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 25

  GottschlingD. E.AparicioO. M.BillingtonB. L.ZakianV. A. (1990). Position effect at S. cerevisiae telomeres: reversible repression of Pol II transcription. Cell63, 751–762. 10.1016/0092-8674(90)90141-Z

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 26

  GreiderC. W. (2012). Molecular biology. Wnt regulates TERT – putting the horse before the cart. Science336, 1519–1520. 10.1126/science.1223785

  • CrossRef
  • Google Scholar

• 27

  GreiderC. W.BlackburnE. H. (1987). The telomere terminal transferase of Tetrahymena is a ribonucleoprotein enzyme with two kinds of primer specificity. Cell51, 887–898. 10.1016/0092-8674(87)90576-9

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 28

  GreiderC. W.BlackburnE. H. (1989). A telomeric sequence in the RNA of Tetrahymena telomerase required for telomere repeat synthesis. Nature337, 331–337. 10.1038/337331a0

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 29

  GriffithJ. D.ComeauL.RosenfieldS.StanselR. M.BianchiA.MossH.et al (1999). Mammalian telomeres end in a large duplex loop. Cell97, 503–514. 10.1016/S0092-8674(00)80760-6

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 30

  GrillariJ.KatingerH.VoglauerR. (2006). Aging and the ubiquitinome: traditional and non-traditional functions of ubiquitin in aging cells and tissues. Exp. Gerontol.41, 1067–1079. 10.1016/j.exger.2006.07.003

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 31

  GugerK. A.GumbinerB. M. (2000). A mode of regulation of beta-catenin signaling activity in Xenopus embryos independent of its levels. Dev. Biol.223, 441–448. 10.1006/dbio.2000.9770

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 32

  HamadaF.BienzM. (2004). The APC tumor suppressor binds to C-terminal binding protein to divert nuclear beta-catenin from TCF. Dev. Cell7, 677–685. 10.1016/j.devcel.2004.08.022

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 33

  HartM. J.De Los SantosR.AlbertI. N.RubinfeldB.PolakisP. (1998). Downregulation of beta-catenin by human Axin and its association with the APC tumor suppressor, beta-catenin and GSK3 beta. Curr. Biol.8, 573–581. 10.1016/S0960-9822(98)70226-X

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 34

  HemannM. T.GreiderC. W. (1999). G-strand overhangs on telomeres in telomerase-deficient mouse cells. Nucleic Acids Res.27, 3964–3969. 10.1093/nar/27.20.3964

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 35

  HendersonB. R. (2000). Nuclear-cytoplasmic shuttling of APC regulates beta-catenin subcellular localization and turnover. Nat. Cell Biol.2, 653–660. 10.1038/35046558

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 36

  HilgerM.MannM. (2012). Triple SILAC to determine stimulus specific interactions in the Wnt pathway. J. Proteome Res.11, 982–994. 10.1021/pr200740a

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 37

  HoffmeyerK.RaggioliA.RudloffS.AntonR.HierholzerA.Del ValleI.et al (2012). Wnt/beta-catenin signaling regulates telomerase in stem cells and cancer cells. Science336, 1549–1554. 10.1126/science.1218370

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 38

  JhoE. H.ZhangT.DomonC.JooC. K.FreundJ. N.CostantiniF. (2002). Wnt/beta-catenin/Tcf signaling induces the transcription of Axin2, a negative regulator of the signaling pathway. Mol. Cell. Biol.22, 1172–1183. 10.1128/MCB.22.4.1172-1183.2002

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 39

  KorinekV.BarkerN.WillertK.MolenaarM.RooseJ.WagenaarG.et al (1998). Two members of the Tcf family implicated in Wnt/beta-catenin signaling during embryogenesis in the mouse. Mol. Cell. Biol.18, 1248–1256.

  • Pubmed Abstract
  • Google Scholar

• 40

  LevisR.HazelriggT.RubinG. M. (1985). Effects of genomic position on the expression of transduced copies of the white gene of Drosophila. Science229, 558–561. 10.1126/science.2992080

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 41

  LiV. S.NgS. S.BoersemaP. J.LowT. Y.KarthausW. R.GerlachJ. P.et al (2012). Wnt signaling through inhibition of beta-catenin degradation in an intact Axin1 complex. Cell149, 1245–1256. 10.1016/j.cell.2012.05.002

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 42

  LiuC.LiY.SemenovM.HanC.BaegG. H.TanY.et al (2002). Control of beta-catenin phosphorylation/degradation by a dual-kinase mechanism. Cell108, 837–847. 10.1016/S0092-8674(02)00685-2

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 43

  LiuJ. P. (1999). Studies of the molecular mechanisms in the regulation of telomerase activity. FASEB J.13, 2091–2104.

  • Pubmed Abstract
  • Google Scholar

• 44

  LiuY.SnowB. E.HandeM. P.YeungD.ErdmannN. J.WakehamA.et al (2000). The telomerase reverse transcriptase is limiting and necessary for telomerase function in vivo. Curr. Biol.10, 1459–1462. 10.1016/S0960-9822(00)00805-8

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 45

  LongheseM. P. (2008). DNA damage response at functional and dysfunctional telomeres. Genes Dev.22, 125–140. 10.1101/gad.1626908

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 46

  LyashenkoN.WinterM.MiglioriniD.BiecheleT.MoonR. T.HartmannC. (2011). Differential requirement for the dual functions of beta-catenin in embryonic stem cell self-renewal and germ layer formation. Nat. Cell Biol.13, 753–761. 10.1038/ncb2260

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 47

  MakarovV. L.LejnineS.BedoyanJ.LangmoreJ. P. (1993). Nucleosomal organization of telomere-specific chromatin in rat. Cell73, 775–787. 10.1016/0092-8674(93)90256-P

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 48

  MartinezP.BlascoM. A. (2011). Telomeric and extra-telomeric roles for telomerase and the telomere-binding proteins. Nat. Rev. Cancer11, 161–176. 10.1038/nrc3025

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 49

  MartinezP.ThanasoulaM.CarlosA. R.Gomez-LopezG.TejeraA. M.SchoeftnerS.et al (2010). Mammalian Rap1 controls telomere function and gene expression through binding to telomeric and extratelomeric sites. Nat. Cell Biol.12, 768–780. 10.1038/ncb2081

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 50

  MolenaarM.Van De WeteringM.OosterwegelM.Peterson-MaduroJ.GodsaveS.KorinekV.et al (1996). XTcf-3 transcription factor mediates beta-catenin-induced axis formation in Xenopus embryos. Cell86, 391–399. 10.1016/S0092-8674(00)80112-9

  • CrossRef
  • Google Scholar

• 51

  MosimannC.HausmannG.BaslerK. (2009). Beta-catenin hits chromatin: regulation of Wnt target gene activation. Nat. Rev. Mol. Cell Biol.10, 276–286. 10.1038/nrm2654

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 52

  NimmoE. R.CranstonG.AllshireR. C. (1994). Telomere-associated chromosome breakage in fission yeast results in variegated expression of adjacent genes. EMBO J.13, 3801–3811.

  • Pubmed Abstract
  • Google Scholar

• 53

  OzawaM.BaribaultH.KemlerR. (1989). The cytoplasmic domain of the cell adhesion molecule uvomorulin associates with three independent proteins structurally related in different species. EMBO J.8, 1711–1717.

  • Pubmed Abstract
  • Google Scholar

• 54

  ParkJ. I.VenteicherA. S.HongJ. Y.ChoiJ.JunS.ShkreliM.et al (2009). Telomerase modulates Wnt signalling by association with target gene chromatin. Nature460, 66–72. 10.1038/nature08137

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 55

  PucciF.GardanoL.HarringtonL. (2013). Short telomeres in ESCs lead to unstable differentiation. Cell Stem Cell12, 479–486. 10.1016/j.stem.2013.01.018

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 56

  RaoA. S.KremenevskajaN.ReschJ.BrabantG. (2005a). Lithium stimulates proliferation in cultured thyrocytes by activating Wnt/beta-catenin signalling. Eur. J. Endocrinol.153, 929–938. 10.1530/eje.1.02038

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 57

  RaoR.ZhangM. Z.ZhaoM.CaiH.HarrisR. C.BreyerM. D.et al (2005b). Lithium treatment inhibits renal GSK-3 activity and promotes cyclooxygenase 2-dependent polyuria. Am. J. Physiol. Renal Physiol.288, F642–F649. 10.1152/ajprenal.00287.2004

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 58

  RedonS.ReichenbachP.LingnerJ. (2010). The non-coding RNA TERRA is a natural ligand and direct inhibitor of human telomerase. Nucleic Acids Res.38, 5797–5806. 10.1093/nar/gkq296

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 59

  RubinfeldB.SouzaB.AlbertI.MullerO.ChamberlainS. H.MasiarzF. R.et al (1993). Association of the APC gene product with beta-catenin. Science262, 1731–1734. 10.1126/science.8259518

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 60

  SamperE.FloresJ. M.BlascoM. A. (2001). Restoration of telomerase activity rescues chromosomal instability and premature aging in Terc^-/- mice with short telomeres. EMBO Rep.2, 800–807. 10.1093/embo-reports/kve174

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 61

  SchoeftnerS.BlascoM. A. (2008). Developmentally regulated transcription of mammalian telomeres by DNA-dependent RNA polymerase II. Nat. Cell Biol.10, 228–236. 10.1038/ncb1685

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 62

  SchoeftnerS.BlascoM. A. (2010). Chromatin regulation and non-coding RNAs at mammalian telomeres. Semin. Cell Dev. Biol.21, 186–193. 10.1016/j.semcdb.2009.09.015

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 63

  SfeirA.de LangeT. (2012). Removal of shelterin reveals the telomere end-protection problem. Science336, 593–597. 10.1126/science.1218498

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 64

  SfeirA. J.ChaiW.ShayJ. W.WrightW. E. (2005). Telomere-end processing the terminal nucleotides of human chromosomes. Mol. Cell18, 131–138. 10.1016/j.molcel.2005.02.035

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 65

  Shippen-LentzD.BlackburnE. H. (1990). Functional evidence for an RNA template in telomerase. Science247, 546–552. 10.1126/science.1689074

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 66

  SierraJ.YoshidaT.JoazeiroC. A.JonesK. A. (2006). The APC tumor suppressor counteracts beta-catenin activation and H3K4 methylation at Wnt target genes. Genes Dev.20, 586–600. 10.1101/gad.1385806

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 67

  SternJ. L.BryanT. M. (2008). Telomerase recruitment to telomeres. Cytogenet. Genome Res.122, 243–254. 10.1159/000167810

  • CrossRef
  • Google Scholar

• 68

  StrongM. A.Vidal-CardenasS. L.KarimB.YuH.GuoN.GreiderC. W. (2011). Phenotypes in mTERT± and mTERT^-/- mice are due to short telomeres, not telomere-independent functions of TERT. Mol. Cell. Biol.31, 2369–2371. 10.1128/MCB.05312-11

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 69

  SutherlandC.LeightonI. A.CohenP. (1993). Inactivation of glycogen synthase kinase-3 beta by phosphorylation: new kinase connections in insulin and growth-factor signalling. Biochem. J.296(Pt 1), 15–19.

  • Pubmed Abstract
  • Google Scholar

• 70

  ValentaT.HausmannG.BaslerK. (2012). The many faces and functions of beta-catenin. EMBO J.31, 2714–2736. 10.1038/emboj.2012.150

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 71

  Vidal-CardenasS. L.GreiderC. W. (2010). Comparing effects of mTR and mTERT deletion on gene expression and DNA damage response: a critical examination of telomere length maintenance-independent roles of telomerase. Nucleic Acids Res.38, 60–71. 10.1093/nar/gkp855

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 72

  WangJ.XieL. Y.AllanS.BeachD.HannonG. J. (1998). Myc activates telomerase. Genes Dev.12, 1769–1774. 10.1101/gad.12.12.1769

  • CrossRef
  • Google Scholar

• 73

  WrightW. E.TesmerV. M.HuffmanK. E.LeveneS. D.ShayJ. W. (1997). Normal human chromosomes have long G-rich telomeric overhangs at one end. Genes Dev.11, 2801–2809. 10.1101/gad.11.21.2801

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 74

  WuP.TakaiH.De LangeT. (2012). Telomeric 3’ overhangs derive from resection by Exo1 and Apollo and fill-in by POT1b-associated CST. Cell150, 39–52. 10.1016/j.cell.2012.05.026

  • Pubmed Abstract
  • CrossRef
  • Google Scholar

• 75

  YamadaS.PokuttaS.DreesF.WeisW. I.NelsonW. J. (2005). Deconstructing the cadherin-catenin-actin complex. Cell123, 889–901. 10.1016/j.cell.2005.09.020

  • Pubmed Abstract
  • CrossRef
  • Google Scholar